Rodents, specifically rats and mice, comprise more than 80% of the animals used in biomedical research, teaching, and testing. The technology of advanced physiological monitoring in rats, mice and other animals is rapidly advancing. (Goode, R. L. G., Klein, H. J., 2002, Miniaturization: an overview of biotechnologies for monitoring the physiology and pathophysiology or rodent animal models, ILAR J., 43:136-46) Researchers are gathering reliable data from a variety of physiological systems in various rodent species using laser Doppler flowmetry, digital sonomicrometry, bioelectrical impedance, and microdialysis, using noninvasive physiological monitoring methods, in instances where the animal is mobile.
Basic research in fields such as neuroscience, physiology, pharmacology, virology, immunology, and oncology use large numbers of rodents to asses the effects of biological and pharmacologically active agents. (Cunliffe-Beamer, T. L., 1993, Applying principles of aseptic surgery to rodents, AWIC Newslett 4:3-6; National Research Council, 1996, Guide for the care and use of laboratory animals, 7th Ed., Washington D.C., National Academy Press) Many of these studies use vascular infusion technology to derive samples for assessing activity, biodistribution, and plasma duration. A variety of vascular infusion and intravascular delivery systems have been used in rodents for many years (Fox, C. E., Beazley, R. M., 1975, Chronic venous catheterization: a technique for implanting and maintaining venous catheters in rats, J. Surg. Res. 18:607-10).
Physiologic monitoring of animals often requires externalization of percutaneous, subcutaneous, or otherwise indwelling catheters or cannulas which are coupled to tethering tubes which are further coupled to stationary infusion pumps, power consoles or meters. Initially, the mobility of the animals was restricted either physically or chemically to prevent entanglement and disruption of the tethering tubes. To achieve a more normal, stress-free environment, swivels were developed as an intermediate device to couple the tethering tube with the pump. The swivel allowed a singly-housed animal to move freely about the cage. Line tangling was prevented by virtue of the swivels.
For withdrawing fluids from animals, a tethering apparatus is essentially the only option for long-term continuous access to an animal for physiological monitoring. (Loughnane, M., Jacobson, A., 2004, Tethered infusion and withdrawal in laboratory animals, Animal Lab News, September/October) (http://animallab.com/articles.asp?pid=73) Tethering systems allow for mobility of animals that are chronically catheterized. (U.S. Pat. No. 4,900,313) One tethering system that has been described consists of a jacket made of a light weight, breathable nylon netting material, a light-weight, highly flexible stainless steel attaching tether, and a cage-mounted swivel to which the tether is anchored (Chatham, A. K, 1985, Jacket and Swivel Tethering Systems, Lab Animal 14(8): 29-33).
Early pioneers in the area of tethered infusion systems incorporated various “home-brew” systems (Guillery, E. N., Chodak, G. W., 1984, A technique for continuous infusion in the mouse bladder, J. Urol. 131:1005-7; Hagmuller, K., et al., 1992, A tail-artery cannulation method for the study of blood parameters in freely moving rats, J. Pharmacol. Toxicol. Methods, 28: 79-83; Hodge, D, Shalev, M, 1992, Dual cannulation: a method for continuous interveneous infusion and repeated blood sampling in unrestrained mice, Lab Anim. Sci., 42: 320-22; Patijn G. A., et al., 1998, Method for continuous infusion into the portal vein of mice, Lab Anim. Sci., 48:379-83). A key element of any tether system is the swivel that allows for rotational movement of the tethered animal. In the late 1960s, Michael Loughnane, a biomedical engineer at Temple University, began to design and build swivels to meet the needs of research investigators for tethered infusion in rats. His continued efforts in this specialty area led to the commercialization of the swivel and many other well-engineered components and systems for tethered infusion and sampling, which are available from Instech Solomon (Plymouth Meeting, Pa.).
The primary factors for selecting the components of an infusion system include study length, study end point, access site, need for blood withdrawal, and periodic versus continuous access. (Nolan, T. E., Klein, H. J., 2002, Methods in vascular infusion biotechnology in research with rodents, ILAR J., 43(3):175-82.) “Acute” studies can last up to one day in duration, while “chronic” studies can last longer than one day, during which time the animal is maintained in a normal physiological state. The need for chronic access invokes a different set of component requirements compared with an acute study. Tethering of the test animal is used when chronic access to a conduit is required in a freely moving animal.
Tethered infusion, which involves continuous intravenous infusion or fluid withdrawal of small laboratory animals, e.g. rats and mice, are the most common applications of the devices and system of the invention. (FIG. 1) Details of the use of catheters, tether attachment devices, swivels, harnesses and infusion pumps are well known in the art (e.g. Instech Solomon).
Tethered infusion systems typically include a subcutaneous button, jacket or harness, tail cuff, or a head block apparatus placed near the catheter exit site. This restraint part of the system connects to a spring tether, which is attached to a swivel mounted to the animal's cage. Button tethers and head blocks require surgical placement and are used for long-term studies. They are fabricated of stainless steel, plastic, Dacron mesh, or silicone and are surgically implanted directly beneath the skin of the animal. Fixation of head blocks requires the use of dental acrylic, which attaches the device to the skull bones.
A saddle, jacket, harness or a button infusion device permits externalization of a catheter or cannula through the scapular region of the animal. (Loughnane, M., & Jacobson, A., 2004, Tethered infusion and withdrawal in laboratory animals, Animal Lab News) (http://animallab.com/articles.asp?pid=73) Jackets and harnesses require no surgical intervention and are commercially available in a number of materials and sizes (e.g., Lomir Biomedical, Inc., Malone, N.Y.; Kent Scientific Corporation, Litchfield, Conn.; Alice King Chatham Medical Arts, Hawthorne, Calif.). Jackets are reusable and are made of cloth or nylon in a vest-like conformation, with two cutouts for the front limbs and a reinforced area over the catheter exit site that attaches to the spring tether. However, because of their cloth construction, jackets are prone to soiling and must be washed or replaced periodically.
Harnesses have been used more recently as alternatives to jackets. A harness, sometimes referred to as a saddle, covers the externalization site of the catheter, and provides a means of tethering or restraining the animal, so that the conduit does not become disengaged from the animal. A button infusion device is made of polyester mesh, metal or polysulfone plastic and is sutured to the fascia under the animal's skin, allowing conduit externalization through the button's center.
The Covance Harness™ (Covance Laboratories, Vienna, Va.) is constructed of a molded elastomer saddle, with attached silicone bands that form a sling around the animal's body. They are commercially available for both rats and mice, are designed to be disposable, and do not require periodic cleaning (Instech Solomon). They present a much smaller contact area to the rodent's skin and are not as likely to interfere with thermoregulation. The saddle serves as the attachment point for the spring tether and as a protective covering for the catheter, catheter exit site, or external terminus of the animal interface, and subsequent couplings.
Stainless steel tubes are inserted through a hole in the Covance Harness for coupling to a catheter emerging from the animal to the swivel extension line. The coupling of the stainless steel tubes to the catheter and then to swivel extension line and subsequent threading of the line through a tether is slow, difficult and cumbersome to attach. Additionally, no quick removal of the system is available; once the animal is connected to the system it cannot be removed from the swivel extension line and catheter without considerable work. This manipulation results in high stress to the animal, which could lead to erroneous experimental results. Minimizing stress to the animal increases reproducibility from animal to animal, reduces animal use, and is ethically justified for animal welfare and for improving study validity. (Nolan, T. E., Klein, H. J., 2002, Methods in vascular infusion biotechnology in research with rodents, ILAR J., 43(3):175-82 (http:/dels.nas.edu/ilar/jour_online/43—3/v4303nolan.asp))
The commercially available vascular infusion technology and accompanying product literature provides ample guidance so that a user can select the optimal system or infusion devices such as catheters and pumps to ensure a successful study outcome when vascular delivery or collection of a sample via the vascular route is desirable. The technologies available for monitoring rodent physiological systems and for monitoring and characterizing specific organ systems such as the cardiovascular and respiratory systems are well known, including methods of vascular access in rodents. (Goode, T. L., Klein, H. J., 2002, Miniaturization: An overview of biotechnologies for monitoring the physiology and pathophysiology of rodent animal models, ILAR J., 43:136-46; Hartley, C. J., et. al., 2002, Noninvasive cardiovascular phenotyping in mice, ILAR J., 43:147-58; Hedlund, L. W., Johnson, G. A., 2002, Mechanical ventilation for imaging the small animal lung, ILAR J., 43:159-74; Nolan, T. E., Klein, H. J., 2002, Methods in vascular infusion biotechnology in research with rodents, ILAR J., 43:175-82).